This work was supported by the division of intramural research at the National Institutes of Health/NHLBI (1ZIAHL006126) and a Lenfant Biomedical Postdoctoral Fellowship awarded to DAL.

1. Preparation of Materials Required to Explant Third Instar Larval Brains
<ol>
	<li>Prepare tools required for microdissection.
	<ol>
		<li>Forge two dissecting tools (a scalpel and a hook) by first placing a single dissecting pin into each pin holder. Secure the pins tightly by hand or with pliers (Figure 1C).</li>
		<li>Use a pair of old forceps to bend one dissecting pin to an approximately 120&#176; angle. Do this under a dissecting microscope to ensure that the top half of the pin lies flat when the pin holder is held in hand. This tool will be used as a scalpel to hold down or slice through tissue.</li>
		<li>Use a pair of old forceps to bend the second pin into a hook that will be used to hold and scrape away tissue.</li>
		<li>Cover each dissecting tool with a pipette tip to protect the tools from damage. With care, well-formed tools may be used for dissection indefinitely. Replace damaged or deformed pins as necessary.</li>
	</ol>
	</li>
	<li>Prepare dissecting medium.
	<ol>
		<li>In a tissue culture hood, dilute the antibiotic-antimycotic supplement into the Schneider&#8217;s culturing medium. Swirl the media to incorporate the supplement.</li>
		<li>Prepare several 5 ml aliquots of the supplemented media. Store all media at 4 &#176;C until ready for use.</li>
		<li>Warm a 5 ml aliquot of supplemented medium to room temperature. Draw the medium into a sterile 5 ml syringe and screw on a sterile 0.2 &#181;m syringe filter.</li>
	</ol>
	</li>
	<li>Select larvae for dissection. Use a dissecting probe to select crawling third instar larvae as the brain will be easiest to dissect. Optional: deposit larvae on a grape agar plate. 
	Note: Do not use overly crowded vials or bottles with &#8220;wet food,&#8221; as these tend to force second or first instar larvae to crawl out prematurely. 
	Note: Some genetic backgrounds will necessitate using younger larvae. This does not affect the protocol, but does make the dissection more difficult.</li>
</ol>
2. Dissection of the Central Nervous System
<ol>
	<li>On a single glass slide, create two 50 ml pools of warm dissecting media separated by about 3 cm. One pool will be used for gross dissection and other used for fine dissection (Figure 1C).</li>
	<li>Transfer several larvae to one of the media drops.</li>
	<li>Move the coverslip with larvae to a dissecting microscope. Zoom into the pool containing the larvae such that the anterior and posterior ends of the larvae are discernible.</li>
	<li>Use a pairs of forceps to grasp the mid-region of a single larva. Take a second pair of forceps in the other hand and grasp the larva close to the same region. Pull the two pairs of forceps apart to gently tear the larva in half. Repeat the above two steps for all larvae on the coverslip.</li>
	<li>Dispose of the posterior halves of the larval body by transferring them to a tissue.</li>
	<li>Zoom into a larva so that the anterior mouthhooks are clearly visible. Use both pairs of forceps to create a small incision, or notch, on opposite sides of the larval mouthhooks between the third thoracic and first anterior denticle bands.</li>
	<li>Grasp the mouthhooks with the forceps in the non-dominant hand. Use the forceps in the other hand to gently peel away cuticle in an anterior-to-posterior direction. Continue to slowly peel away cuticle until the central nervous system is exposed.</li>
	<li>Isolate the central nervous system from the rest of the larva by tearing tissues with the forceps. Use caution not to disturb the central nervous system. 
	Critical step: Do not tug on the central nervous system! Discard damaged samples with a torn ventral nerve cord (Figure 2A) or distorted optic lobes (Figures 2B and 2B&#8217;).</li>
	<li>Repeat the above two steps for all larvae on the coverslip. Transfer the healthy explanted tissue to the second (clean) pool of medium on the coverslip for the fine dissection stage.</li>
	<li>Use the dissecting pin tools to clear away the peripheral tissues (e.g., eye, wing, and leg discs) from the brain. Slide the scalpel between the wanted (brain) and the unwanted tissue, pinning the connective tissue to the coverglass. Use the hook to gently tease away the undesired tissue while simultaneously pressing the scalpel to the coverslip in saw-like movements.</li>
	<li>Critical step: Work slowly and deliberately to remove all discs from each brain. Use caution not to disturb the brain morphology. A healthy brain will have an intact ventral nerve cord and symmetric, round optic lobes (Figure 2C).</li>
</ol>
3. Mounting Explanted Brains for Live Imaging
<ol>
	<li>Invert a 50 mm gas-permeable culture dish with the clear film facing up (Figure 2D). Depress the syringe to deposit a small drop (about 30 &#181;l) of medium onto the center of the dish.</li>
	<li>Transfer the isolated brains, one at a time, to the drop of medium on the dish. Use the hook tool to scoop up a brain underneath the optic lobes. Alternatively, use forceps to grasp axons projecting from the ventral nerve cord.</li>
	<li>Collect 5-10 brains in the drop of medium. Submerge the brains by using the dissecting pin tools to push individual brains to the bottom of the drop of medium as many of them will be trapped at the meniscus (Figure 2E).</li>
	<li>Orient the brains using the dissecting pin tools to align brains depending on the NBs to be imaged (Figure 3A). To image the dorsal NBs, place the ventral nerve cord facing down (along the membrane). To image the anterio-ventral NBs, place the brains with the ventral nerve cord facing upward (away from membrane). Align the brains such that all the ventral nerve cords point in the same direction within the x-y plane.</li>
	<li>Use a syringe or plastic transfer pipette to place 4 halocarbon (HC) oil drops (about 30 &#181;l each) onto the gas permeable membrane. The volume of each drop of oil should be equivalent to each other and to the drop of medium. Space the drops of oil to correspond to the four corners of a coverslip centered around the drop of culturing medium. Once complete, the five drops (four drops of oil and one drop of medium in the center) will resemble the five facets on Western-style dice (Figure 3B).</li>
	<li>Lower a #1.5 22 mm coverslip on the assembly such that it hits all 5 drops at the same time. Maintaining an even pressure across the samples reduces the likelihood that they will be disrupted. Wait about 3 min as the oil and medium disperse under the weight of the coverslip. A cross-like shape of media will form (Figures 3C and 3D).</li>
	<li>Critical step: Lower the coverslip onto the surface of the brains by removing excess media using a tissue paper wick (Figure 3C). Do this while watching the sample under the dissecting scope. As media is removed, the coverslip will lower. Stop once the coverslip touches the brains. Do not over-wick as this will cause tissue distortion or brain explosions.</li>
	<li>If imaging dorsal NBs (Figure 3E), move to step 3.9. If imaging anterio-ventral NBs, the coverslip must be slightly moved (by gently nudging it with forceps) in the direction of the ventral nerve cord tips. This causes the brain to rotate, which brings the brain lobes in direct contact with the coverslip to facilitate imaging (Figure 3F).</li>
	<li>Seal the chamber by encircling the coverslip with small amounts of halocarbon oil (Figure 3D). Excess oil oozing from the coverslip should be minimized and blotted away with a tissue.</li>
</ol>
4. Live Imaging of Central Brain Neuroblasts
Note: For this work, a Nikon Eclipse Ti spinning-disk confocal microscope (inverted microscope) was used for live imaging; details of our setup have been previously published31. Alternatively, the sample can be imaged using an upright system.
<ol>
	<li>Add a drop of immersion oil to the coverslip just above the mounted brains. This will help center the objective directly above the sample.</li>
	<li>Critical step: Maintain brains at a constant temperature of 25 &#176;C with a stage incubator. Gently position the mounted sample into the stage incubator and cover the chamber with its lid.</li>
	<li>Locate central brain NBs using transmitted light by focusing on the large, round cells that reside in the central and medial areas of the optic lobes; do not use epi-fluorescence. This becomes easier with practice. It is very important to expose the sample to the least amount of light as possible. Do not waste time imaging brains that are damaged.</li>
	<li>Once in place, take quick exposures using the confocal to determine proper location and depth. Limit depth to the first 10-15 &#181;m. Adjust as needed, and take another test image.
	<ol>
		<li>Optional step: Collect a test movie of the ventral nerve cord to ensure there is no x-y drift. If drift is detected, wait 15-30 min for the sample to settle. If the drift does not settle in 30 min, prepare a new sample. Major drift can usually be traced back to the addition of excess oil, or the unequal size of oil drops during mounting.</li>
	</ol>
	</li>
	<li>To eliminate axial drift (z-drift) use a continuous automatic focusing device that uses an infrared LED. Alternatively, manually correct the focal plane.</li>
	<li>Once a NB of interest is identified, proceed with imaging regimen. As with all live cell imaging, the amount of laser light, exposure time, camera binning, objective magnification, time and/or z-step interval must all be empirically optimized for a particular experiment to answer the question at hand. The goal is to minimize light damage while still collecting useful data. See discussion for additional guidance.
	<ol>
		<li>Image the entire volume of the NB by collecting 12-14 images spaced by 1 &#181;m in the z-axis. Adjust the time resolution to capture single or multiple cell cycles of the same NB. As a general guideline, use 10-30 sec intervals for single cell cycle imaging and 1-3 min intervals for multiple cell cycles.</li>
	</ol>
	</li>
	<li>Critical step: Confirm that the sample is healthy by monitoring the duration of mitosis. If mitosis takes more than 15 min, move on to another brain. A healthy brain will show many NBs within the same field of view undergoing several rounds of mitosis. Note that each cell cycle is slightly longer than the previous one due to light damage.</li>
	<li>Once imaging is complete, disassemble the incubation chamber, and discard everything but the gas-permeable culturing dishes. Rinse the culture dish surface with 95% ethanol and wipe well with a tissue. Repeat two more times.</li>
</ol>
5. Preparation of Brains for Immunofluorescence
<ol>
	<li>Collect 20 or more explanted brains into a 1.5 ml tube containing approximately 0.2 ml of supplemented medium.</li>
	<li>Remove medium from the tube and rinse brains once with 0.5 ml PBSTx (Phosphate Buffered Saline, PBS, with 0.3% Triton X-100). Let brains settle to the bottom of the tube between all exchanges of liquid. The Triton X-100 detergent is used to permeabilize the tissue. Rinsing typically takes less than 30 sec; simply remove the PBSTx once brains have settled to the bottom of the tube.</li>
	<li>Replace the solution with 0.5 ml of 9% electron microscopy grade paraformaldehyde diluted in PBSTx. Incubate with nutation for 15 min at room temperature.</li>
	<li>Dispose of fixative according to hazardous waste regulations.</li>
	<li>Wash the samples with 0.5 ml PBSTx for 15 min. Repeat the wash step with fresh PBSTx two more times.</li>
	<li>Replace the solution with 0.5 ml PBT (PBS,1% Bovine Serum Albumin (BSA), and 0.1% Tween-20). Incubate with nutation for 1 hr at room temperature. This step blocks the nonspecific binding of antibody to the tissue.</li>
	<li>Replace the solution with 0.5 ml primary antibody diluted in PBT supplemented with 4% normal goat serum (NGS). Incubate with nutation overnight at 4 &#176;C.</li>
	<li>Discard or save the diluted primary antibody.</li>
	<li>Wash the samples with 0.5 ml PBT for 15 min. Repeat the wash step with fresh PBT two more times.</li>
	<li>Replace the solution with 0.5 ml of modified PBT (PBS, 2% BSA, 0.1% Tween-20, and 4% NGS). Incubate with nutation for 1 hr at room temperature.</li>
	<li>Replace the solution with 0.5 ml secondary antibody diluted into modified PBT. Incubate with nutation for 2 hr at room temperature.</li>
	<li>Replace the solution with 0.5 ml PBST (PBS with 0.1% Tween-20). Incubate with nutation for 15 min at room temperature. Repeat the wash step with fresh PBST two more times.</li>
</ol>
6. Mounting Samples for Immunofluorescence
<ol>
	<li>Carefully transfer the samples as a drop on a glass slide, restricting the drop towards the left of the center of slide.</li>
	<li>Add a large drop (about 2 cm in diameter) of mounting medium (for example, Aqua-Poly/Mount) to the center of the slide. Avoid the pool of PBST that is toward the left.</li>
	<li>Use forceps to transfer the brains from the pool of PBST to the pool of mounting medium. Adding mounting medium directly on top of the brains may disrupt their orientation and should be avoided.</li>
	<li>Under a light microscope, arrange the samples in the mounting medium with the side that will be imaged facing up.</li>
	<li>With the slide under the light microscope, slowly lower a #1.5 glass coverslip on top of the samples. Optional step: Dislodge minor air bubbles by lightly pushing on the coverslip.</li>
	<li>Allow the mounting medium to polymerize for several hours, or overnight, by lying slides flat, light-protected, at room temperature.</li>
	<li>Slides may be imaged the next day, or stored prior to imaging.</li>
</ol>

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J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+protocol+for+culturing+Drosophila+melanogaster+stage+9+egg+chambers+for+live+imaging.">A protocol for culturing Drosophila melanogaster stage 9 egg chambers for live imaging.</a> Nat. Protoc. 2, 2467-2473 (2007).</li><li>Rusan, N. M., Peifer, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+role+for+a+novel+centrosome+cycle+in+asymmetric+cell+division.">A role for a novel centrosome cycle in asymmetric cell division.</a> J. Cell Biol. 177, 13-20 (2007).</li><li>Martinez-Campos, M., Basto, R., Baker, J., Kernan, M., Raff, J. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+Drosophila+pericentrin-like+protein+is+essential+for+cilia/flagella+function,+but+appears+to+be+dispensable+for+mitosis.">The Drosophila pericentrin-like protein is essential for cilia/flagella function, but appears to be dispensable for mitosis.</a> J. Cell Biol. 165, 673-683 (2004).</li><li>Dix, C. I., Raff, J. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Drosophila+Spd-2+recruits+PCM+to+the+sperm+centriole,+but+is+dispensable+for+centriole+duplication.">Drosophila Spd-2 recruits PCM to the sperm centriole, but is dispensable for centriole duplication.</a> Curr. Biol. 17, 1759-1764 (2007).</li><li>Moutinho-Santos, T., Sampaio, P., Amorim, I., Costa, M., Sunkel, C. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vivo+localisation+of+the+mitotic+POLO+kinase+shows+a+highly+dynamic+association+with+the+mitotic+apparatus+during+early+embryogenesis+in+Drosophila.">In vivo localisation of the mitotic POLO kinase shows a highly dynamic association with the mitotic apparatus during early embryogenesis in Drosophila.</a> Biol. Cell. 91, 585-596 (1999).</li><li>Megraw, T. L., Kilaru, S., Turner, F. R., Kaufman, T. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+centrosome+is+a+dynamic+structure+that+ejects+PCM+flares.">The centrosome is a dynamic structure that ejects PCM flares.</a> J. Cell Sci. 115, 4707-4718 (2002).</li></ol>

The authors have nothing to disclose.

Live cell imaging is an invaluable approach used to study the mechanisms that regulate ACD of stem cells, the differentiation of cell lineages, and the morphogenesis of complex tissues. Although research groups have historically employed a variety of techniques to visualize NB ACD, these approaches can be generally grouped into two categories that primarily differ with respect to whether the brain tissue is left intact or dissociated.
Imaging dissociated NBs has its advantages. For one, dissoc...

Stem cells maintain a balance of differentiation and self-renewal to generate cellular diversity during early development and replace damaged cells in adult tissues. Regulation of this homeostasis prevents both the loss and over-expansion of the stem cell population, which can result in deleterious tissue degeneration or tumorigenesis.
Neuroblasts (NBs) are the widely studied neural stem cells of the Drosophila central nervous system (Figure 1A) that first form during embryonic stages1,2. NBs undergo repeated rounds of asymmetric cell division (ACD) to produce two unequally fated cells: a self-renewing stem cell and a differentiating cell. ACD is guided by centrosomes, the non-membranous organelles that act as the microtubule-organizing centers of most cells3. During mitosis, NB centrosomes organize and orient the bipolar mitotic spindle along the apical-basal polarity axis. Upon cleavage of the dividing NB, apical fate determinants that specify the stem cell fate, and basal fate determinants that specify differentiation, are segregated into the unequal daughter cells.
In the larval central brain, two types of NBs may be distinguished by their number, position, transcription factor expression, and cell lineage (Figure 1A)4-6. Type I NBs are the most abundant, and about 90 of them populate the anterior and posterior sides of each optic lobe of the brain7. These NBs express the transcription factor Asense (Ase), and they characteristically divide into one self-renewing NB and one smaller ganglion mother cell (GMC; Figure 1B). Each GMC undergoes a single terminal division to generate two neurons or glia (Figure 1B). In contrast, the eight Type II NBs that populate the posterior side of each optic lobe lack Ase expression5. They undergo ACD to produce one self-renewing NB and one intermediate neural progenitor (INP).
The INP, in turn, divides asymmetrically three to five times. Each of these divisions results in regeneration of the INP and the production of a single GMC4. Collectively, the specific NB identity and the temporal order of GMC birth gives rise to the astounding neuronal diversity of the adult central nervous system.
Understanding the cell biology that underlies NB ACD has been vastly improved through the use of live cell imaging techniques. Published protocols used by researchers to image live NBs vary widely. Overall, however, these methods may be grouped into two general categories distinguished by whether the larval brain is left intact or mechanically dissociated. Both techniques have distinct advantages and disadvantages depending on the researcher&#8217;s application.
Early reports revealing live NB cell divisions involve some degree of manual dissociation of the larval brain. These protocols detail smearing8 or teasing apart9 the brain in order to grow short-term primary cultures of the NBs on glass coverslips. To improve imaging, the round NBs are usually flattened on the coverslips with either glass slides8 or agarose pads9. Although flattened cells have improved optics, these techniques often lead to NB mitotic defects, including regression of the cleavage furrow and the inability to divide more than one time. Therefore, protocols that involve both manual dissociation and physical distortion of the larval brain tissue are generally only suited to very short-term (i.e., one cell cycle or less) applications. Likewise, semi-squashing or completely flattening intact brains between glass slides and coverslips limits the time one may spend imaging a given specimen to an approximately 30 min period10. Despite this limitation, this approach has been used successfully11-14.
Recent efforts to image live dissociated NBs limit physical distortion of the isolated cells. These techniques are particularly useful for applications where it is necessary to sustain cell cultures for multiple cell division cycles. For example, dispersed cells from manually dissociated brains may be partially embedded in a clot made from the mixture of fibrinogen and thrombin15 for long-term imaging16. Alternatively, explanted brains may be first chemically dissociated with the enzyme collagenase, next manually disrupted by repeated passage through a pipet tip, and subsequently plated on poly-L-lysine-coated glass bottom dishes17,18. Coating glass dishes with either fibrinogen or poly-L-lysine promotes the attachment and slight spreading of round cells, bringing them as close to the coverslip as possible without major physical distortion. In addition, these methods involve culturing the NBs in medium that may be readily exchanged for pharmacological perturbation experiments18. Overall, directly plating dispersed NBs improves imaging optics without sacrificing long-term imaging capabilities. Recently, a protocol describing the long-term culturing of dissociated NBs has been detailed19.
However, this approach is not without its limitations. There are several caveats to imaging live dissociated NBs. In a field of dispersed cells, for example, it is difficult to identify specific NB lineages that are spatially organized in the intact brain1. Likewise, distinguishing the Type I versus Type II NBs within dissociated tissue is also challenging without the expression of lineage tracers or subtype-specific transgenes, such as worniu-GAL4, asense-GAL8020. Although these transgenes are quite useful for some applications, they do limit researchers to those transgenes expressed under the control of the UAS enhancer element21 and require more complex genetic schemes. Studies that concern the spatial or temporal development of NBs, therefore, require in vivo imaging in the context of an intact tissue.
Moreover, studies of dissociated embryonic NBs indicate that the physical contact of adjacent cells is critical for the interphase localization of key polarity determinants22,23. These studies show completely isolated NBs no longer maintain the invariant mitotic spindle axis. Instead, these cells display a more randomized spindle axis and wide angles of separation between successive GMC buds, perhaps due to the loss of apical centrosome positioning22. Although the relationship between the larval NBs and their neighboring cells has not been extensively studied, it is worth considering that the larval stem cell microenvironment may impinge upon cell polarity or other behaviors. To maintain the physiological context of larval NBs, we image ACD from whole brains.
Given the inherent limitations associated with imaging dissociated NBs, our lab and others have established protocols to image successive rounds of NB ACD from whole larval brains. Techniques to image intact brains often replace the rigid surface of glass on one side of the culture chamber with a flexible membrane to prevent tissue distortion or damage and allow for gas exchange. Some methods involve mounting explanted brains in a mixture of insect culturing medium, serum, and ascorbic acid with the fat bodies isolated from ten larvae24. The isolated fat bodies are added because they secrete a mitogen known to stimulate NB cell division25. This protocol preserves the integrity of the tissue for over 3&#160;hr&#160;and is, therefore, amenable to long-term imaging24,26-28. Recently, a protocol describing the long-term imaging of intact larval brains in the presence of ascorbic acid, serum, and fat bodies has been detailed29. Importantly, because the tissue is neither exposed nor immobilized, this approach is less useful for applications where culturing medium is exchanged (e.g., drug studies, immunodepletion, etc.).
Our lab has simplified the whole mount preparation of live larval brains for long-term imaging30,31. The main advantage to our protocol over others is its simplicity: our observations indicate neither serum, ascorbic acid, insulin, nor fat bodies are necessary additives to image successive rounds of NB ACD. Although additives, such as insulin, have been useful for the prolonged imaging of stem cell divisions32 and collective cell migration in Drosophila ovaries33, they appear to be dispensable for NB ACD, as our imaging medium consists of a simple mixture of standard insect cell culturing medium supplemented with antibiotic-antimycotic to prevent contamination. Through the use of reusable gas-permeable or glass bottom culture dishes, we have been able to sustain several rounds of successive NB ACDs. The same explanted samples can readily be used for immunofluorescence and other procedures with fixed tissue. In this protocol, we detail the dissection of intact larval brains, as well as the preparation of brains for both live and fixed analysis.

<table border="0" cellpadding="0" cellspacing="0" width="851">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:380px;">Lumox Tissue Culture Dish 50X12 mm</td>
			<td style="width:185px;">Sarstedt</td>
			<td style="width:119px;">94.6077.410</td>
			<td style="width:167px;">A reusable culture dish with a gas-permeable and optically clear film base.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:380px;">Schneider&#39;s S2 Medium</td>
			<td style="width:185px;">Invitrogen</td>
			<td style="width:119px;">21720-024</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:380px;">Gibco Antibiotic-Antimycotic (100x)</td>
			<td style="width:185px;">Invitrogen</td>
			<td style="width:119px;">15240-062</td>
			<td>A supplement containing penicillin (10,000 U/mL), streptomycin (10,000 mg/mL), and amphotericin B (25 mg/mL) that is added to Schneider&#39;s S2 media to prevent bacterial and fungal contamination.&#160;</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:380px;">Glass coverslips, #1.5 22X22 mm</td>
			<td style="width:185px;">Fisher Scientific</td>
			<td style="width:119px;">12-541-B</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:380px;">Halocarbon oil 700</td>
			<td style="width:185px;">Sigma-Aldrich</td>
			<td style="width:119px;">H8898</td>
			<td>A high viscosity inert oil that is clear and colorless.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:380px;">pair of nickel-plated pin holders, 17 cm long</td>
			<td style="width:185px;">Fine Science Tools</td>
			<td style="width:119px;">26018-17</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:380px;">Minutien dissecting pins</td>
			<td style="width:185px;">Fine Science Tools</td>
			<td style="width:119px;">26002-10</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:380px;">pair of Dumont #5 Forceps</td>
			<td style="width:185px;">Fine Science Tools</td>
			<td style="width:119px;">11252-20</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:380px;">Dissecting needle/probe with plastic handle</td>
			<td style="width:185px;">Fisher Scientific</td>
			<td style="width:119px;">08-965-A</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:380px;">Syringe filter, 0.22 mm pore size</td>
			<td style="width:185px;">Fisher Scientific</td>
			<td style="width:119px;">09-719A</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:380px;">Syringe 5 mL&#160;</td>
			<td style="width:185px;">BD Biosciences</td>
			<td style="width:119px;">309646</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:380px;">plastic transfer pipet</td>
			<td style="width:185px;">Fisher Scientific</td>
			<td style="width:119px;">S30467-1</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:380px;">paraformaldehyde, 32% EM-grade</td>
			<td style="width:185px;">Electron Microscopy Sciences</td>
			<td style="width:119px;">15714</td>
			<td>CAUTION: Formaldehyde is toxic; it should be handled in a fume hood. Follow MSDS guidelines for safe handling.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:380px;">DAPI (4&#39;, 6-diamidino-2-phenylindole, dihydrochloride)</td>
			<td style="width:185px;">Invitrogen</td>
			<td style="width:119px;">D1306</td>
			<td>A dye that labels DNA and is excited by UV light.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:380px;">Aqua/PolyMount; Polysciences, Inc.</td>
			<td style="width:185px;">Fisher Scientific</td>
			<td style="width:119px;">18606-20</td>
			<td>A water-soluble mounting medium.</td>
		</tr>
	</tbody>
</table>

live imaging of drosophila larval neuroblasts

Stem cells divide asymmetrically to generate two progeny cells with unequal fate potential: a self-renewing stem cell and a differentiating cell. Given their relevance to development and disease, understanding the mechanisms that govern asymmetric stem cell division has been a robust area of study. Because they are genetically tractable and undergo successive rounds of cell division about once every hour, the stem cells of the Drosophila central nervous system, or neuroblasts, are indispensable models for the study of stem cell division. About 100 neural stem cells are located near the surface of each of the two larval brain lobes, making this model system particularly useful for live imaging microscopy studies. In this work, we review several approaches widely used to visualize stem cell divisions, and we address the relative advantages and disadvantages of those techniques that employ dissociated versus intact brain tissues. We also detail our simplified protocol used to explant whole brains from third instar larvae for live cell imaging and fixed analysis applications.

Live imaging has elucidated a number of mechanisms that regulate NB ACD. Prior to image acquisition, it is necessary to determine the optimal imaging conditions. It is necessary to balance the frame capture rate with the duration of live cell imaging. For fine cellular analysis, for example, increased temporal and optical resolution may be required. When visualizing a single NB cell cycle (Figure 4A), the rate at which images are captured may be increased without introducing photodamage. Typically, singl...

Watch this Scientific Journal Video about Live Imaging of Drosophila Larval Neuroblasts at JoVE.com

Live Imaging of Drosophila Larval Neuroblasts

This protocol details a streamlined method used to conduct live cell imaging in the context of an intact larval brain. Live cell imaging approaches are invaluable for the study of asymmetric neural stem cell divisions as well as other neurogenic and developmental processes, consistently uncovering mechanisms that were previously overlooked.

Live Imaging of Drosophila Larval Neuroblasts

Stem cells divide asymmetrically to generate two progeny cells with unequal fate potential: a self-renewing stem cell and a differentiating cell. Given ...

live-imaging-of-drosophila-larval-neuroblasts

Research

JoVE Journal

Neuroscience

14.7K Görüntüleme.  National Institutes of Health.  Bu protokol, dokunulmamış larva beyin bağlamında canlı hücre görüntüleme yapmak için kullanılan akıcı bir yöntem göstermektedir. Canlı hücre görüntüleme yaklaşımları sürekli daha önce gözardı mekanizmaları ortaya çıkarılması, asimetrik nöral kök hücre bölünmeleri çalışmanın yanı sıra diğer nörolojik ve gelişimsel süreçleri için paha biçilmezdir. 

Video: Live Imaging of Drosophila Larval Neuroblasts

Magnetic Resonance Spectroscopy of live Drosophila melanogaster using Magic Angle Spinning

High-Resolution Magic Angle Spinning (HRMAS) proton magnetic resonance spectroscopy (1H-MRS) is a novel non-destructive technique that improves spectral line-widths and allows high-resolution spectra to be obtained from extracts, intact cells, cell cultures, and more importantly intact tissue to investigate relationships between metabolites and cellular processes. In vivo HRMAS 1H-MRS studies have yet to be reported in the live fruit fly Drosophila melanogaster. Drosophila, as a simpler genetic organism, allows the multiple biological functions and various evolutionarily conserved signaling pathways to be examined at the whole organism level and it is a useful model for investigating genetics and physiology. To this end, we developed and implemented an in vivo HRMAS 1H-MRS method to investigate live Drosophila at 14.1 T. Here, we outline an HRMAS 1H-MRS protocol for the molecular characterization of Drosophila with a conventional MR spectrometer equipped with an HRMAS probe. This technique is a novel, in vivo, non-destructive Drosophila metabolite measurement approach, which enables the identification of disease biomarkers and thus may contribute to novel therapeutic development.

Magnetic Resonance Spectroscopy of live Drosophila melanogaster using Magic Angle Spinning

This technique enables the use of high-resolution magic angle spinning proton MR spectroscopy (HRMAS 1H-MRS) for molecular characterization of live Drosophila melanogaster with a conventional 14.1 tesla spectrometer equipped with an HRMAS probe.

High-Resolution Magic Angle Spinning (HRMAS) proton magnetic resonance spectroscopy (1H-MRS) is a novel non-destructive technique that improves spectral ...

Visualization of Larval Segmental Nerves in 3rd Instar Drosophila Larval Preparations

Drosophila melanogaster is emerging as a powerful model system for studying the development and function of the nervous system, particularly because of its convenient genetics and fully sequenced genome. Additionally, the larval nervous system is an ideal model system to study mechanisms of axonal transport as the larval segmental nerves contain bundles of axons with their cell bodies located within the brain and their nerve terminals ending along the length of the body. Here we describe the procedure for visualization of synaptic vesicle proteins within larval segmental nerves. If done correctly, all components of the nervous system, along with associated tissues such as muscles and NMJs, remain intact, undamaged, and ready to be visualized. 3rd instar larvae carrying various mutations are dissected, fixed, incubated with synaptic vesicle antibodies, visualized and compared to wild type larvae. This procedure can be adapted for several different synaptic or neuronal antibodies and changes in the distribution of a variety of proteins can be easily observed within larval segmental nerves.

Visualization of Larval Segmental Nerves in 3rd Instar Drosophila Larval Preparations

Drosophila melanogaster larvae provide an ideal model system to investigate the mechanisms of axonal transport within larval segmental nerves. Using this procedure, 3rd instar larvae carrying various mutations can be compared to wild type larvae.

Drosophila melanogaster is emerging as a powerful model system for studying the development and function of the nervous system, particularly because of ...

In vivo Visualization of Synaptic Vesicles Within Drosophila Larval Segmental Axons

Elucidating the mechanisms of axonal transport has shown to be very important in determining how defects in long distance transport affect different neurological diseases. Defects in this essential process can have detrimental effects on neuronal functioning and development. We have developed a dissection protocol that is designed to expose the Drosophila larval segmental nerves to view axonal transport in real time. We have adapted this protocol for live imaging from the one published by Hurd and Saxton (1996) used for immunolocalizatin of larval segmental nerves. Careful dissection and proper buffer conditions are critical for maximizing the lifespan of the dissected larvae. When properly done, dissected larvae have shown robust vesicle transport for 2-3 hours under physiological conditions. We use the UAS-GAL4 method 1 to express GFP-tagged APP or synaptotagmin vesicles within a single axon or many axons in larval segmental nerves by using different neuronal GAL4 drivers. Other fluorescently tagged markers, for example mitochrondria (MitoTracker) or lysosomes (LysoTracker), can be also applied to the larvae before viewing. GFP-vesicle movement and particle movement can be viewed simultaneously using separate wavelengths.

In vivo Visualization of Synaptic Vesicles Within Drosophila Larval Segmental Axons

This protocol discusses the live dissection of Drosophila larvae for the purpose of imaging the movement of GFP tagged axonal vesicles on microtubule tracks.

Elucidating the mechanisms of axonal transport has shown to be very important in determining how defects in long distance transport affect different ...

Live-cell Imaging of Sensory Organ Precursor Cells in Intact Drosophila Pupae

Since the discovery of Green Fluorescent Protein (GFP), there has been a revolutionary change in the use of live-cell imaging as a tool for understanding fundamental biological mechanisms. Striking progress has been particularly evident in Drosophila, whose extensive toolkit of mutants and transgenic lines provides a convenient model to study evolutionarily-conserved developmental and cell biological mechanisms. We are interested in understanding the mechanisms that control cell fate specification in the adult peripheral nervous system (PNS) in Drosophila. Bristles that cover the head, thorax, abdomen, legs and wings of the adult fly are individual mechanosensory organs, and have been studied as a model system for understanding mechanisms of Notch-dependent cell fate decisions. Sensory organ precursor (SOP) cells of the microchaetes (or small bristles), are distributed throughout the epithelium of the pupal thorax, and are specified during the first 12 hours after the onset of pupariation. After specification, the SOP cells begin to divide, segregating the cell fate determinant Numb to one daughter cell during mitosis. Numb functions as a cell-autonomous inhibitor of the Notch signaling pathway.
Here, we show a method to follow protein dynamics in SOP cell and its progeny within the intact pupal thorax using a combination of tissue-specific Gal4 drivers and GFP-tagged fusion proteins 1,2.This technique has the advantage over fixed tissue or cultured explants because it allows us to follow the entire development of an organ from specification of the neural precursor to growth and terminal differentiation of the organ. We can therefore directly correlate changes in cell behavior to changes in terminal differentiation. Moreover, we can combine the live imaging technique with mosaic analysis with a repressible cell marker (MARCM) system to assess the dynamics of tagged proteins in mitotic SOPs under mutant or wildtype conditions. Using this technique, we and others have revealed novel insights into regulation of asymmetric cell division and the control of Notch signaling activation in SOP cells (examples include references 1-6,7 ,8).

Live-cell Imaging of Sensory Organ Precursor Cells in Intact Drosophila Pupae

In this video, we describe a method for live cell imaging of asymmetrically dividing sensory organ progenitor cells and epidermal cells in intact Drosophila pupae

Since the discovery of Green Fluorescent Protein (GFP), there has been a revolutionary change in the use of live-cell imaging as a tool for understanding ...

Dissection and Immunohistochemistry of Larval, Pupal and Adult Drosophila Retinas

The compound eye of Drosophila melanogaster consists of about 750 ommatidia (unit eyes). Each ommatidium is composed of about 20 cells, including lens-secreting cone cells, pigment cells, a bristle cell and eight photoreceptors (PRs) R1-R8 2. The PRs have specialized microvillar structures, the rhabdomeres, which contain light-sensitive pigments, the Rhodopsins (Rhs). The rhabdomeres of six PRs (R1-R6) form a trapezoid and contain Rh1 3 4. The rhabdomeres of R7 and R8 are positioned in tandem in the center of the trapezoid and share the same path of light. R7 and R8 PRs stochastically express different combinations of Rhs in two main subtypes5: In the 'p' subtype, Rh3 in pR7s is coupled with Rh5 in pR8s, whereas in the 'y' subtype, Rh4 in yR7s is associated with Rh6 in yR8s 6 7 8.
Early specification of PRs and development of ommatidia begins in the larval eye-antennal imaginal disc, a monolayer of epithelial cells. A wave of differentiation sweeps across the disc9 and initiates the assembly of undifferentiated cells into ommatidia10-11. The 'founder cell' R8 is specified first and recruits R1-6 and then R7 12-14. Subsequently, during pupal development, PR differentiation leads to extensive morphological changes 15, including rhabdomere formation, synaptogenesis and eventually rh expression.
In this protocol, we describe methods for retinal dissections and immunohistochemistry at three defined periods of retina development, which can be applied to address a variety of questions concerning retinal formation and developmental pathways. Here, we use these methods to visualize the stepwise PR differentiation at the single-cell level in whole mount larval, midpupal and adult retinas (Figure 1).

Dissection and Immunohistochemistry of Larval, Pupal and Adult Drosophila Retinas

The Drosophila retina is a crystal-like lattice composed of a small number of cell types that are generated in a stereotyped manner 1. Its amenability to sophisticated genetic analysis allows the study of complex developmental programs. This protocol describes dissections and immunohistochemistry of retinas at three discrete developmental stages, with a focus on photoreceptor differentiation.

The compound eye of Drosophila melanogaster consists of about 750 ommatidia (unit eyes). Each ommatidium is composed of about 20 cells, including ...

Functional Analysis of the Larval Feeding Circuit in Drosophila

The serotonergic feeding circuit in Drosophila melanogaster larvae can be used to investigate neuronal substrates of critical importance during the development of the circuit. Using the functional output of the circuit, feeding, changes in the neuronal architecture of the stomatogastric system can be visualized. Feeding behavior can be recorded by observing the rate of retraction of the mouth hooks, which receive innervation from the brain. Locomotor behavior is used as a physiological control for feeding, since larvae use their mouth hooks to traverse across an agar substrate. Changes in feeding behavior can be correlated with the axonal architecture of the neurites innervating the gut. Using immunohistochemistry it is possible to visualize and quantitate these changes. Improper handling of the larvae during behavior paradigms can alter data as they are very sensitive to manipulations. Proper imaging of the neurite architecture innervating the gut is critical for precise quantitation of number and size of varicosities as well as the extent of branch nodes. Analysis of most circuits allow only for visualization of neurite architecture or behavioral effects; however, this model allows one to correlate the functional output of the circuit with the impairments in neuronal architecture.

Functional Analysis of the Larval Feeding Circuit in Drosophila

The feeding circuit in Drosophila melanogaster larvae serves a simple yet powerful model that allows changes in feeding rate to be correlated with alterations in the stomatogastric neural circuitry. This circuit is composed of central serotonergic neurons that send projections to the mouth hooks as well as the foregut.

The serotonergic  feeding circuit in Drosophila  melanogaster larvae can be used to investigate neuronal substrates of  critical importance during the ...

Why Quantification Matters: Characterization of Phenotypes at the Drosophila Larval Neuromuscular Junction

Most studies on morphogenesis rely on qualitative descriptions of how anatomical traits are affected by the disruption of specific genes and genetic pathways. Quantitative descriptions are rarely performed, although genetic manipulations produce a range of phenotypic effects and variations are observed even among individuals within control groups.&#160;Emerging evidence shows that morphology, size and location of organelles play a previously underappreciated, yet fundamental role in cell function and survival. Here we provide step-by-step instructions for performing quantitative analyses of phenotypes at the Drosophila larval neuromuscular junction (NMJ). We use several reliable immuno-histochemical markers combined with bio-imaging techniques and morphometric analyses to examine the effects of genetic mutations on specific cellular processes. In particular, we focus on the quantitative analysis of phenotypes affecting morphology, size and position of nuclei within the striated muscles of Drosophila larvae. The Drosophila larval NMJ is a valuable experimental model to investigate the molecular mechanisms underlying the structure and the function of the neuromuscular system, both in health and disease. However, the methodologies we describe here can be extended to other systems as well.

Why Quantification Matters: Characterization of Phenotypes at the Drosophila Larval Neuromuscular Junction

Morphology, size and location of intracellular organelles are evolutionarily conserved and appear to directly affect their function. Understanding the molecular mechanisms underlying these processes has become an important goal of modern biology. Here we show how these studies can be facilitated by the application of quantitative techniques.

Most studies on morphogenesis rely on qualitative descriptions of how anatomical traits are affected by the disruption of specific genes and genetic ...

Using Linear Agarose Channels to Study Drosophila Larval Crawling Behavior

Drosophila larval crawling is emerging as a powerful model to study neural control of sensorimotor behavior. However, larval crawling behavior on flat open surfaces is complex,&#160;including: pausing, turning, and meandering. This complexity in the repertoire of movement hinders detailed analysis of the events occurring during a single crawl stride cycle. To overcome this obstacle, linear agarose channels were made that constrain larval behavior to straight, sustained, rhythmic crawling. In principle, because agarose channels and the Drosophila larval body are both optically clear, the movement of larval structures labeled by genetically-encoded fluorescent probes can be monitored in intact, freely-moving larvae. In the past, larvae were placed in linear channels and crawling at the level of whole organism, segment, and muscle were analyzed1. In the future, larvae crawling in channels can be used for calcium imaging to monitor neuronal activity. Moreover, these methods can be used with larvae of any genotype and with any researcher-designed channel. Thus the protocol presented below is widely applicable for studies using the Drosophila larva as a model to understand motor control.

Using Linear Agarose Channels to Study Drosophila Larval Crawling Behavior

The Drosophila larva is a powerful model system to study neural control of behavior. This publication describes the use of linear agarose channels to elicit sustained bouts of linear crawling and methods to quantify the dynamics of larval structures during repetitive crawling behavior.

Drosophila larval crawling is emerging as a powerful model to study neural control of sensorimotor behavior. However, larval crawling behavior on flat ...

Single-cell Resolution Fluorescence Live Imaging of Drosophila Circadian Clocks in Larval Brain Culture

The circadian pacemaker circuit orchestrates rhythmic behavioral and physiological outputs coordinated with environmental cues, such as day/night cycles. The molecular clock within each pacemaker neuron generates circadian rhythms in gene expression, which underlie the rhythmic neuronal functions essential to the operation of the circuit. Investigation of the properties of the individual molecular oscillators in different subclasses of pacemaker neurons and their interaction with neuronal signaling yields a better understanding of the circadian pacemaker circuit. Here, we present a time-lapse fluorescent microscopy approach developed to monitor the molecular clockwork in clock neurons of cultured Drosophila larval brain. This method allows the multi-day recording of the rhythms of genetically encoded fluorescent circadian reporters at single-cell resolution. This setup can be combined with pharmacological manipulations to closely analyze real-time response of the molecular clock to various compounds. Beyond circadian rhythms, this multipurpose method in combination with powerful Drosophila genetic techniques offers the possibility to study diverse neuronal or molecular processes in live brain tissue.

Single-cell Resolution Fluorescence Live Imaging of Drosophila Circadian Clocks in Larval Brain Culture

The goal of this protocol is to establish ex vivo Drosophila larval brain culture optimized to monitor circadian molecular rhythms with long-term fluorescence time-lapse imaging. The application of this method to pharmacological assays is also discussed.

The circadian pacemaker circuit orchestrates rhythmic behavioral and physiological outputs coordinated with environmental cues, such as day/night cycles. ...

Tracking Drosophila Larval Behavior in Response to Optogenetic Stimulation of Olfactory Neurons

The ability of insects to navigate toward odor sources is based on the activities of their first-order olfactory receptor neurons (ORNs). While a considerable amount of information has been generated regarding ORN responses to odorants, the role of specific ORNs in driving behavioral responses remains poorly understood. Complications in behavior analyses arise due to different volatilities of odorants that activate individual ORNs, multiple ORNs activated by single odorants, and the difficulty in replicating naturally observed temporal variations in olfactory stimuli using conventional odor-delivery methods in the laboratory. Here, we describe a protocol that analyzes Drosophila larval behavior in response to simultaneous optogenetic stimulation of its ORNs. The optogenetic technology used here allows for specificity of ORN activation and precise control of temporal patterns of ORN activation. Corresponding larval movement is tracked, digitally recorded, and analyzed using custom written software. By replacing odor stimuli with light stimuli, this method allows for a more precise control of individual ORN activation in order to study its impact on larval behavior. Our method could be further extended to study the impact of second-order projection neurons (PNs) as well as local neurons (LNs) on larval behavior. This method will thus enable a comprehensive dissection of olfactory circuit function and complement studies on how olfactory neuron activities translate in to behavior responses.

Tracking Drosophila Larval Behavior in Response to Optogenetic Stimulation of Olfactory Neurons

This protocol analyzes navigational behavior of Drosophila larva in response to simultaneous optogenetic stimulation of its olfactory neurons. Light of 630 nm wavelength is used to activate individual olfactory neurons expressing a red-shifted channel rhodopsin. Larval movement is simultaneously tracked, digitally recorded, and analyzed using custom-written software.